Plasmodium vivax infection compromises reticulocyte stability.

doi:10.21203/rs.3.rs-53799/v1

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Plasmodium vivax infection compromises reticulocyte stability.

https://doi.org/10.21203/rs.3.rs-53799/v1

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published 11 Mar, 2021

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The structural integrity of the host red blood cell (RBC) must be maintained for propagation of Plasmodium spp. during the disease causing blood-stage of malaria infection. Plasmodium vivax infection is restricted to reticulocytes. To assess the impact of P. vivax infection on reticulocyte stability, we developed a flow cytometry-based assay capable of measuring osmotic stability within heterogeneous RBC populations. We found that P. vivax preferred young reticulocytes are more osmotically stable than older reticulocytes and normocytes, and P. vivax infection decreased reticulocyte stability to levels observed for RBC disorders that cause hemolytic anemia. Moreover, P. vivax reticulocyte destabilization was more significant than P. falciparum normocyte destabilization. Finally, we found that P. vivax new permeability pathways contribute to the decreased osmotic stability of infected-reticulocytes. These results reveal a key vulnerability of P. vivax that could be manipulated to yield both in vitro culture and novel therapeutics.

General Cell Biology & Physiology

General Microbiology

Parasitology

Plasmodium vivax

reticulocyte stability

malaria

RBC

The unique biconcave shape and structural dynamics of red blood cells (RBCs) permit traversal of the circulatory system and the delivery of oxygen to tissues, and are determined by molecules that make up the RBC cytoskeleton and membrane transporters ¹. Naturally occurring polymorphisms in molecules such as PIEZO1 and Ankyrin-1, that disrupt cell volume regulation and cytoskeleton, are associated with the clinically-relevant RBC phenotypes of hereditary xerocytosis (HX) ² and hereditary spherocytosis (HS), ³ respectively. These mutations manifest in structurally compromised RBCs with abnormal shapes (stomatocytes and spherocytes) and premature hemolysis in vivo ⁴. A feature of these mutant RBCs is abnormal sensitivity to hypotonic challenge, which is decreased in HX and increased in HS.

During the Plasmodium spp. intra-erythrocytic developmental cycle (IDC), parasite remodeling alters the physiology of the host RBC. In the case of the most well studied of these parasites, P. falciparum, changes include disruption of the cytoskeleton by parasite proteins inserted into the RBC membrane ⁵ and increased plasma membrane permeability ^6,7 that culminate in the infected RBC taking on a spherical shape ⁸. These changes compromise the structural integrity of the host RBC, and consequently decrease the osmotic stability of P. falciparum-infected RBCs ^6,7,9, while increasing the propensity of infected cells to be destroyed in circulation via clearance by the spleen as well as extravascular hemolysis ¹⁰.

Plasmodium vivax, the most globally widespread of the malaria parasites infecting humans ¹¹, exhibits a strict tropism for reticulocytes, the youngest of circulating RBCs ^12–14. This is in contrast to P. falciparum, which also infects the mature normocyte RBC fraction. The absence of an in vitro culture system and appropriate experimental methods have impeded the study of how P. vivax impacts the structural integrity of the host reticulocyte. The reticulocyte compartment itself is highly heterogeneous due to the dynamic final steps of RBC maturation, which involves enucleation, continued synthesis of hemoglobin, loss of membrane and organelles, and reconfiguration of the cytoskeleton ^15–23. All P. falciparum studies have been done in the older, more abundant normocyte fraction. Furthermore, recent observations that P. vivax increases reticulocyte deformability ^13,24,25 while P. falciparum decreases normocyte deformability ^5,24,26 suggest that the pathophysiology of P. vivax reticulocyte infection are fundamentally different than P. falciparum normocyte infection.

Characterizing RBC stability under osmotic stress is a powerful method for probing the structural integrity of the cytoskeleton and membrane transporter activity, and therefore is invaluable for assessing RBC disorders that compromise in vivo fitness such as HS ^2–4. Reticulocytes ^27–29 and erythroid progenitors ³⁰ are more resistant to hypotonic lysis than normocytes, but the dynamics of osmotic stability during erythropoiesis and reticulocyte maturation are unknown. Osmotic stability is traditionally assessed by measuring release of hemoglobin from bulk RBC preparations in increasingly hypotonic solutions. This methodology cannot resolve the osmotic stability of specific RBC subpopulations within heterogeneous RBC preparations. Furthermore, using free hemoglobin as the read-out for RBC osmotic stability is problematic as hemoglobin levels are in flux in both parasite-infected RBCs and uninfected reticulocytes, because Plasmodium spp. digest hemoglobin as they mature ³¹, and reticulocytes are actively synthesizing hemoglobin ³². Flow cytometry, with its ability to examine discrete cell subsets within heterogeneous populations, has the potential to better define RBC osmotic stability. Previous studies have demonstrated the capacity of flow cytometry to detect hemolysis ^33–36, but are not amendable to collecting sufficient data on rare cell populations.

Here we have developed a flow cytometric osmotic stability assay that quantifies the RBCs that survive hypotonic lysis, and we have used it to assess the osmotic stability dynamics of reticulocytes and erythroid progenitors and the impact of P. vivax infection on reticulocyte osmotic stability. We found that osmotic stability steadily decreased during erythroid differentiation and reticulocyte maturation. Of enucleated RBCs the youngest CD71 + reticulocytes, preferred by P. vivax, are the most osmotically stable, but upon infection with P. vivax were destabilized to levels observed for RBCs from individuals with hemolytic anemia. Furthermore, P. vivax-infected reticulocytes were significantly less stable than P. falciparum-infected normocytes. Finally, we found that the decreased stability of P. vivax-infected reticulocytes corresponded with the appearance of P. vivax new permeability pathways (NPPs). In summary, by establishing an osmotic stability assay capable of assessing RBCs at the single cell level, we have been able to define the osmotic stability dynamics within heterogeneous reticulocyte and malaria-infected RBC populations. The observation that reticulocyte osmotic stability is reduced by P. vivax infection reveals a key vulnerability of the parasite.

Flow cytometry analysis of RBC osmotic stability. While flow cytometry has been employed to study osmotic stability, previous studies have not harnessed the capacity of flow cytometry to analyze multiple cell populations within a single specimen ^33–36. Therefore we developed a flow cytometric osmotic stability assay that yields the same hemolysis curves as the traditional hemoglobin absorbance assay and is capable of defining osmotic stability dynamics within heterogeneous RBC populations. Hemolysis curves were generated by first stopping lysis of RBCs in increasingly hypotonic solutions with a lysis quenching solution consisting of PBS and cell counting beads (Fig. 1A). This analysis revealed typical RBC forward scatter (FSC) and side scatter (SSC) profiles for non-lytic isotonic conditions, and the subsequent appearance of a low-FSC/low-SSC population in hypotonic, lytic conditions (Fig. 1B and Supplemental Fig. 1).

Hypothesizing that the low-FSC/low-SSC population was RBC ghosts, fluorescent phalloidin, which is excluded from intact cells but binds actin in the cytoskeleton of permeabilized cells was included in the osmotic stability assay. We found that phalloidin stained the FSC-low population that appears in lytic conditions, indicating that the flow lysis assay is sensitive to RBC ghosts (Fig. 1C and 1D). The FSC-low/SSC-low RBC ghost population was subsequently excluded when quantifying RBC osmotic stability. With this strategy the lysis curves produced by the flow lysis assay were indistinguishable from those of the hemoglobin absorbance lysis assay (Fig. 1E). No difference in the lysis₅₀ values (mOsm at which 50% of RBCs lyse) obtained for normal RBCs using the two lysis assays confirmed the accuracy of the flow lysis assay. To test the limits of the flow lysis assay, we measured the osmotic stability of hereditary xerocytosis (HX) RBCs that are resistant to hypotonic lysis. With HX RBCs as well, we observed no difference in lysis₅₀ values generated by the two assays (Fig. 1F).

Finally, as P. vivax osmotic stability studies included cryopreserved samples and were performed on RBCs maintained in vitro, we examined the effect of (i) cold storage and (ii) P. vivax in vitro culture conditions on RBC osmotic stability. As expected ^37,38, we observed variation in the osmotic stability of normal RBCs taken from twelve different donors with lysis₅₀ values ranging from 93.9 to 137.9 mOsm (variation of 11.2%) (Fig. 1G). Subsequent storage of RBCs from six different donors at 4ºC for two weeks resulted in no appreciable change (mean slope − 0.063 ±0.24 SEM, Pearson r = 0.16) in osmotic stability (Fig. 1H). However, when cryopreserved RBCs and RBCs stored at 4ºC were transferred to P. vivax in vitro culture conditions, 24 hours later osmotic stability increased by 13% ±2.6 and 14.6% ±0.8 (Fig. 1I).

Osmotic stability decreases during erythroid differentiation and reticulocyte maturation.

Having established the capacity of flow cytometry to measure RBC osmotic stability, we next examined the osmotic stability dynamics within the RBC fractions that harbor P. vivax infection. To this end, we took advantage of the capacity of flow cytometry to track discrete cell populations, to assess the osmotic stability of nucleated RBC precursors, reticulocytes, and normocytes in bone marrow aspirates (Fig. 2A and Supplemental 2A). We found that fluorescent labeling of reticulocytes and nucleated RBC precursors present within bone marrow aspirates allowed us to quantitate the osmotic stability dynamics of each of these RBC subpopulations simultaneously (Fig. 2B). For nucleated RBC precursors we additionally found that lysed cells were also identifiable with a live/dead stain (Supplemental 2B). Importantly, as both osmotic stability studies with bone marrow aspirates and clinical P. vivax samples relied on Percoll enrichment to raise reticulocytemia and P. vivax parasitemia to reliably measurable levels, we found that Percoll had no effect on the osmotic stability of reticulocytes and nucleated precursors from bone marrow aspirates (Supplemental 2C).

The lysis₅₀ values obtained for RBC precursors and reticulocytes from bone marrow aspirates revealed that nucleated RBC precursors (DNA + CD71+) were the most osmotically stable (Lysis₅₀ 70.1 ±5.8) followed by the youngest CD71 + RNA + DNA- reticulocytes (Lysis₅₀ 87.3 ±8.0), older CD71- RNA + DNA- reticulocytes (Lysis₅₀ 107.4 ±7.3) and CD71- RNA- DNA- normocytes (Lysis₅₀ 114.5 ±5.4) (Fig. 2B and 2C). To establish osmotic stability dynamics during erythropoiesis, we assessed the osmotic stability of erythroid progenitors differentiated in vitro from CD34 + stem cells ^28,39. This study revealed that osmotic stability decreased as cells progressed in vitro from majority basophilic and polychromatic normoblasts at day 9 (Lysis₅₀ 73.8 ±3.6) to a majority polychromatic and orthochromatic normoblast population at day 11 (Lysis₅₀ 102.7 ±15.6), and then further decreased as cells progressed through final orthochromatic normoblast maturation occurring between day 14 (Lysis₅₀ 102.2 ±2.0) day 17 (Lysis₅₀ 126.2 ±11.2), and day 20 (Lysis₅₀ 143.4 ±15.1) (Fig. 2D). Additionally, consistent with a previous study ²⁸, CD71 + reticulocytes generated in vitro (Lysis₅₀ 110.1 ±3.7) were less stable than CD71 + reticulocytes from bone marrow samples (Lysis₅₀ 87.3 ±8.0) (Supplemental Fig. 2D). Together these results demonstrate the capacity of flow cytometry to assess the osmotic stability of discrete RBC subsets within heterogeneous populations and clearly shows that erythroid development and reticulocyte maturation are associated with significant changes in osmotic stability.

P. vivax infection reduces reticulocyte osmotic stability.

Having established that the youngest CD71 + reticulocytes that are preferred by P. vivax for invasion ^13,14 are the most osmotically stable of all enucleated RBCs, we next assessed the impact of P. vivax infection on reticulocyte osmotic stability. In the absence of continuous P. vivax in vitro culture, cryopreserved clinical P. vivax samples are an invaluable resource for investigating P. vivax biology ^40–43. Cognizant of the decreased stability of cryopreserved RBCs, however (Fig. 1I), we first directly compared the in vitro survival and osmotic stability of cryopreserved and non-cryopreserved P. vivax clinical samples as parasites progressed through the IDC (Fig. 3A and B and Supplemental Fig. 3A). Consistent with previous studies ^40–43, we observed a 78% ±3.3 and 63% ±30.8 loss of P. vivax infected-reticulocytes prior to completion of the IDC in vitro (44-hour cultures) for Brazilian cryopreserved and Indian non-cryopreserved clinical samples respectively (Fig. 3C).

Parallel osmotic stability measurements revealed that, as observed previously (Fig. 1I), the stability of cryopreserved uninfected RBC populations increased upon transfer to in vitro culture, while non-cryopreserved uninfected RBCs remained steady during the course of culture (Supplemental Fig. 3B and 3C). For P. vivax-infected reticulocytes, we observed that cryopreserved stage I ring-infected reticulocytes were less stable than non-cryopreserved stage I rings. For subsequent time points we observed (i) no difference in the osmotic stability of cryopreserved and non-cryopreserved P. vivax-infected reticulocytes, and (ii) a decrease in the stability of P. vivax-infected reticulocytes as they progressed through the IDC (Fig. 3D). Moreover, when we consider the clinical laboratory cutoff for normal RBCs (lysis₅₀171 mOsm or 0.5% NaCl), the osmotic stability of both cryopreserved and non-cryopreserved P. vivax-infected reticulocytes in 24- and 44-hour cultures fell into the range of osmotic stabilities associated with hemolytic anemias. Finally, in order to take advantage of the availability of cryopreserved P. vivax clinical samples while also minimizing the influence of cryopreservation, we excluded the 1-hour post thaw time point from subsequent analysis.

We next examined the degree of instability P. vivax induced in the host reticulocyte by comparing the osmotic stability of CD71- and CD71 + P. vivax-infected reticulocytes to that of uninfected CD71 + reticulocytes. Due to reticulocyte maturation ⁴⁴, this analysis was limited to the first 24-hours of in vitro culture, as the frequency of CD71 + P. vivax-infected and uninfected reticulocytes respectively decreased by 56% ±8.0 and 64% ±3.5 between 1 and 24-hours of culture, and then fell below the limit of detection (0.05% CD71 + reticulocytes) between 24 and 44 hours. Of note, the persistence of P. vivax-infected CD71 + reticulocytes in our ex vivo cultures through 24 hours (Supplemental Fig. 3D and 3E) is longer than previously observed for CD71 + cord blood reticulocytes invaded in vitro by P. vivax ¹³. The reason for this discrepancy is not immediately evident and therefore subject for future investigation. We found infected reticulocytes were less stable than uninfected CD71 + reticulocytes at all-time points assessed (8-, 16-, and 24-hour cultures), and the progression of P. vivax through the IDC further decreased reticulocyte osmotic stability with the appearance of stage III late trophozoite forms in 24-hour cultures corresponding to P. vivax-infected CD71 + and CD71- reticulocytes being 71.4% ±12.0 and 74.0% ±14.6 less stable than uninfected CD71 + reticulocytes (Fig. 3D). Finally no difference in the stability of CD71 + and CD71- infected-reticulocytes indicated that P. vivax infection and not reticulocyte age is the primary determinate of the stability of P. vivax-infected reticulocytes (Supplemental Fig. 3F).

P. vivax induces greater host cell instability than P. falciparum

Finally we compared P. vivax destabilization of reticulocytes with P. falciparum destabilization of normocytes. To account for the influence of cryopreservation on our P. vivax osmotic stability studies, we assessed the in vitro survival and osmotic stability of cryopreserved P. falciparum clone 3D7 P2G12 ⁴⁵. We observed similar progression of P. vivax and P. falciparum through the asexual IDC but a greater frequency of sexual gametocyte forms for P. vivax. In vitro survival, however, was markedly different, with a 77.3% ±6.5 survival rate observed for P. falciparum infected-normocytes at 44-hours of culture compared to a 22.4% ±0.03 survival rate for P. vivax infected-reticulocytes (Fig. 3C and 3F).

We subsequently assessed the osmotic stability of uninfected and P. falciparum trophozoite and schizonts stage-infected normocytes in 24- and 44-hour cultures (the points at which P. vivax-infected reticulocytes were most destabilized). This analysis revealed no difference in the osmotic stability of infected (majority trophozoite) and uninfected normocytes in 24-hour cultures and a reduction in infected normocyte (majority schizonts) stability of 23.6% ±6.9 compared to uninfected normocytes in 44 hour cultures (Fig. 3G). This is in contrast to P. vivax, which had decreased reticulocyte stability by 74.0% ±14.6 by the time parasites had matured to the trophozoite form in 24 hour cultures. Furthermore, direct comparison of the osmotic stability of P. falciparum-infected normocytes and P. vivax-infected reticulocytes revealed that P. vivax-infected reticulocytes (max Lysis₅₀ 184.1 ±8.3, 24-hour culture) are significantly less stable, p < 0.0003, than P. falciparum-infected normocytes (max Lysis₅₀ 107.8 ±4.7, 44-hour culture) (Fig. 3H).

Appearance of P. vivax new permeability pathways corresponds with decreased stability of P. vivax-infected reticulocytes.

New permeability pathways (NPPs) in related malaria parasites, P. falciparum and P. knowlesi, increase the permeability of the infected RBC to certain solutes ^6,7. To determine whether P. vivax possesses NPPs that are contributing to the decreased stability of the host cell, we assessed the sensitivity of Percoll-enriched cryopreserved Brazilian clinical P. vivax samples to the NPP antagonists D-sorbitol and L-alanine. We found that uninfected RBCs were not sensitive to D-sorbitol or L-alanine (data not shown). For P. vivax-infected reticulocytes, we found that early stage parasites present in 8-hour cultures were resistant to D-sorbitol and L-alanine lysis, while the appearance of stage III late trophozoite parasites in 16-hour cultures corresponded with P. vivax-infected reticulocytes lysing in isotonic D-sorbitol (16-hour − 47.9% ±9.0, 24-hour − 45.2% ±9.2, and 44-hour − 57.5% ±7.7) and L-alanine solutions (16-hour − 60.7% ±10.5, 24-hour − 61.0% ±9.2, and 44-hour − 55.8% ±2.3). Finally, the NPP inhibitor, furosemide, protected P. vivax-infected reticulocytes from D-sorbitol and L-alanine lysis (Fig. 4A and 4B). The incomplete lysis of P. vivax-infected cells by D-sorbitol and L-alanine along with variation in the sensitivity of different P. vivax clinical isolates to D-sorbitol and L-alanine lysis are potentially driven by (i) the high frequency of more stable gametocytes (42.4% ±9.5 of P. vivax-infected reticulocytes in 44-hour cultures) ^46,47 (Fig. 3C), (ii) variable NPP activity in different P. vivax isolates, or (iii) P. vivax being less sensitive to D-sorbitol lysis than P. falciparum^48,49.

Structural stability of P. vivax-infected reticulocytes, critical for the successful propagation of blood-stage malaria infections, has not been thoroughly assessed due to lack of appropriate methodology compatible with the unique and challenging biology of P. vivax. To study the impact of P. vivax infection on the structural integrity of the host reticulocyte, we adapted the osmotic stability assay to be analyzed by flow cytometry. This permitted us to work with the limited cell numbers and heterogeneity characteristic of both the reticulocyte niche to which P. vivax infection is restricted as well as P. vivax-infected clinical samples. Observations made in the proceeding studies that the youngest CD71 + reticulocyte subset, which P. vivax preferentially invades, was the most osmotically stable and that P. vivax-infected reticulocytes were less stable than uninfected CD71 + reticulocytes, indicate that P. vivax severely compromises the structural integrity of the host reticulocyte. The single-cell resolution of the flow cytometry osmotic stability assay has additional applications including: tracking the impact of erythropoiesis perturbations on overall RBC osmotic stability and examination of osmotic stability dynamics within heterozygous hematological diseases such as sickle cell and G6PD deficiency.

The decreasing osmotic stability we observed in the later stages of erythropoiesis and then continuing through reticulocyte maturation is likely driven by three major changes occurring to erythroid cells during their differentiation: (i) intracellular solutes changes due to changing membrane transporter abundance and activity ^19,21, (ii) cytoskeleton remodeling ^17,22, and (iii) the progressive reduction in the surface to volume ratio driven by membrane loss ²³. Considering previous reports of membrane instability in reticulocytes ⁵⁰, one might hypothesize that reticulocytes would be less osmotically stable than normocytes. The observation of the opposite by us and others ^27–29 indicates that either the factors that determine osmotic stability, such as the higher surface area to volume ratio and greater abundance and activity of membrane transporters in reticulocytes, are more dominant than reticulocyte membrane instability, or that the biology underlying membrane instability and osmotic stability are independent of one another. Ultimately, the factors that determine osmotic stability during erythropoiesis and reticulocyte maturation are likely multifactorial and highly dynamic. Despite the underlying complexity, recent observations of the importance of cholesterol for in vitro differentiated reticulocyte osmotic stability ²⁸ suggest that pursuing the determinants of erythroid progenitor and reticulocyte osmotic stability has the potential to yield strategies for generating more viable RBCs from in vitro cultures.

Our study revealed that P. vivax infection decreased the osmotic stability of the host reticulocyte. Though consistent with what has been observed for other Plasmodium spp. ^6,7,9,51,52, P. vivax decreased reticulocyte osmotic stability to a significantly greater degree than P. falciparum decreased normocyte osmotic stability and to a level on par with RBC instability observed for hemolytic anemias like HS. Interestingly P. knowlesi-infected normocytes have also been reported to be less stable than P. falciparum-infected normocytes⁴⁸. Comparing this data set with ours however, we estimate P. vivax-infected reticulocytes are even less stable (70% less stable than P. falciparum-infected normocytes) than P. knowlesi-infected normocytes (30% less stable than P. falciparum-infected normocytes). These results, specifically the compromised stability of mature P. vivax-infected reticulocytes, are consistent with a study in which the majority of late stage P. vivax-infected reticulocytes lysed when passed through 2 µm microfluidic channels ²⁵ and suggest that the instability induced by P. vivax significantly increases the risk of pre-mature hemolysis of infected reticulocytes in vivo. Considering these findings in the context of the protection from Plasmodium infection afforded by RBC polymorphisms such as HS, thalassemia, and G6PD deficiency, it is possible that a mechanism underlying protection from P. vivax may be premature hemolysis of infected RBCs.

The observation that P. vivax NPPs are associated with reduced stability of host reticulocytes raises several questions including: (i) are P. vivax NPPs required for the acquisition of essential nutrients from the host serum and (ii) what host and or parasite transporters are responsible for the observed NPP activity. In P. falciparum, Clag3.1 and Clag3.2 have been identified as parasite mediators of NPP activity in infected RBCs ^53,54, and P. vivax possesses a clag family of genes ⁵⁵. Despite the aforementioned differences, the very presence of NPP activity in P. vivax-infected reticulocytes suggests some overlap between the underlying factors responsible for the reduced osmotic stability of P. falciparum- and P. vivax-infected cells.

Together our data support a model in which P. vivax-preferred CD71 + reticulocytes are more osmotically stable than CD71- reticulocytes and normocytes, but, upon P. vivax infection and subsequent maturation, the infected host cells undergo a precipitous loss of osmotic stability. Additionally, the onset of NPP activity corresponds with the decreasing osmotic stability of infected reticulocytes (Fig. 4C). Furthermore, the magnitude of instability induced by P. vivax is on par with the RBC instability observed for hemolytic anemias like HS. As a result, P. vivax-infected reticulocytes likely exhibit increased rates of intravascular hemolysis and premature clearance from circulation similar to what is observed for RBCs from individuals with hemolytic anemias. Future studies will focus on the mechanisms underlying these changes in osmotic stability that impact the structural integrity of the host reticulocyte and potentially its ability to survive in vivo. Furthermore, identifying strategies for stabilizing the P. vivax-infected reticulocyte could prove key to culture-adapting P. vivax ⁵⁶. An additionally intriguing focus of future work, is the identification of therapeutic strategies that take advantage of the decreased stability of P. vivax-infected reticulocytes.

Ethics approval:

Anonymized discarded human bone marrow samples negative for blasts or dis-erythropoietic conditions were obtained under Boston Children's Hospital Institutional Review Board (IRB) protocol # 04-02-017R. For Brazilian clinical P. vivax samples, informed consent was obtained from all patients. Study protocols for Brazilian parasite sample collection were approved by the IRB of the Institute of Biomedical Sciences, University of São Paulo, Brazil (1169/CEPSH, 2014). For Indian P. vivax samples, the human subject protocol was approved by the ethics boards at Goa Medical College and Hospital (no number assigned), the University of Washington (42271), and the Division of Microbiology and Infectious Diseases of the National Institutes of Health (11–0074). The overall research program, under which the Indian parasites were collected, was also approved by the Government of India Health Ministry Screening Committee.

Parasite Sample Collection

Brazilian clinical P. vivax samples were collected in the town of Mâncio Lima, Acre State, and processed as described elsewhere ⁵⁷. The collections were performed in the context of a randomized, open-label clinical trial (NCT02691910). Indian P. vivax samples were collected with similar protocols but using CF11 for leukodepletion, as described previously ⁵⁸. This occurred at Goa Medial College and Hospital in Bambolim, Goa, in conjunction with the Malaria Evolution in South Asia International Center of Excellence in Malaria Research and the University of Washington.

Bone marrow and parasite enrichment

Bone marrow aspirates, cryopreserved Brazilian clinical P. vivax samples and the P. falciparum 3D7 P2G12 lab strain were enriched on 1.080 g/mL KCl high Percoll gradients as previously described ⁵⁹. Briefly, 2 mL of re-suspended cells (up to 25% hematocrit) were layered on 3 mL 1.080 g/mL KCl high Percoll gradient and spun for 15 minutes at 1,200 x g. Subsequently, the interface was removed, washed, and applied to assays. For P. falciparum 3D7 P2G12 samples, the interface and pellet were washed and recombined before being applied to assays.

CD34 + in vitro RBC differentiation

CD34 + hematopoetic stem cells (HSCs) purchased commercially (Lonza) were differentiated in vitro following the previously described three-step differentiation protocol ^28,39. Giemsa-stained cytospins were prepared and 200 cells per slide (1000x magnification) were called as either proerythroblast, basophilic normoblast, polychromatic normoblast, orthochromatic normoblast or reticulocyte.

P. vivax and P. falciparum in vitro culture

Indian and Brazilian clinical P. vivax samples and P. falciparum 3D7 P2G12 were cultured at 100 × 10⁶ cells per mL in IMDM (Gibco) containing 10% AB⁺ heat-inactivated sera and 50 µg/mL gentamicin at 37^oC in 5% CO_2, 1% O₂, and 94% N₂ as previously described ⁴⁰. Hemacolor-stained cytospins were prepared and 200 cells per slide (1000x magnification) were called as either asexual stage (I-V) or gametocyte ⁶⁰.

Flow Cytometry. For the described flow cytometry experiments, 1 × 10⁷ cells were washed with flow buffer prior to staining with combinations of the following fluorophores: Thiazole Orange, 1:1000 (Thermo Fisher Scientific), Vybrant DyeCycle Violet, 1:5000 (Thermo Fisher Scientific) α-CD71-APC, 1:25 (Miltenyi Biotech), α-GypA-FITC, 1:100 (Stem Cell Technologies), Vybrant DyeCycle Green, 1:5000 (Thermo Fisher Scientific) and FITC-Phalloidin, 1:200 (Thermo Fisher Scientific). Bone marrow and in vitro cultured RBC samples were stained at 4ºC for 20 minutes and P. vivax and P. falciparum samples at 37ºC for 20 minutes. For bone marrow and in vitro-cultured cell samples, propidium iodide, 1:500 (Thermo Fisher Scientific) was added to samples prior to flow cytometry analysis. All flow cytometry experiments were acquired on a Miltenyi MACSQuant instrument equipped with 405-nm, 488-nm, and 638-nm lasers and data were analyzed using FlowJo (Version 10.4).

Osmotic stability assays. Lysis solutions were made as previously described ⁶¹ and the osmolarity of each was measured with a vapor pressure osmometer (Wescor Vapro 5520). For flow cytometric osmotic stability assays, 1 × 10⁶ cells (stained with appropriate fluorescent dyes and or antibodies) at 5 × 10⁶ cells/mL were incubated in lysis solutions ranging from ~ 300 mOsm to 0 mOsm (distilled water) for 10 minutes at 37ºC. Lysis was stopped with 4x volume of quenching solution (1:10 AccuCheck Counting Beads (Thermo Fisher Scientific) to flow buffer). Only cells with normal flow cytometric FSC/SSC profiles (as defined by the control 300 mOsm condition) were considered to be intact. For assays containing nucleated cells, nuclei and dead cells were excluded from the intact cell population. The ratio of beads to intact cells was used to calculate the absolute number of cells remaining in each lytic condition and percent lysis was determined by normalization to the 300 mOsm control condition (0% lysis). For hemoglobin absorbance osmotic stability assays, cells were lysed as described above before 4x volume of flow buffer was added to stop lysis. Cells were pelleted and the absorbance (380, 415, 450, and 540 nm) of supernatants measured on a Spectramax M5 plate reader. The amount of hemoglobin present in supernatants was calculated by the Harboe method ⁶². Percent lysis was determined by normalization to the 300 mOsm control condition (0% lysis).

RBC ghost assay. For RBC ghost detection by flow cytometry, FITC conjugated phalloidin was added to all lysis and quenching solutions. For RBC ghost detection by immunofluorescence microscopy, RBCs suspended in PBS containing FITC-Phalloidin were allowed to settle on a coverslip before a hypotonic solution (57 mOsm) containing FITC-Phalloidin was added and RBC lysis and ghost formation examined by live video microscopy on a Zeiss AxioObserver.Z1. Images were processed using Fiji software.

Sorbitol and Alanine hemolysis assay. 1 × 10⁶ cells taken from P. vivax cultures were incubated for 30 minutes at 37^oC in flow buffer, 280 mM sorbitol with 1% BSA, or 280 mM alanine with 1% BSA in the presence or absence of 100 µM furosemide. Following incubation, cells were washed twice with flow buffer, and stained for flow cytometry analysis.

Data Sharing Statement. For original data, please contact [email protected].

Author Contributions

MAC UK GWR – Experimental design, execution and data analysis

LC, AM, EG, PKR, CB, MUF – Clinical management of uninfected and infected blood sample collection, including patient management and ethical clearance

MAC UK GWR CB MTD – Data interpretation and manuscript writing

Acknowledgments

This study was supported by U.S. NIH/NHLBI F32 HL136173, NIH R01 AI140751, NIH R01 HL139337 and NIH/NIAID MESA-ICEMR Program Project U19 AI89688, the Canadian Institutes of Health Research Postdoctoral Fellowship, and a Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) research scholarship. The clinical trial that originated samples from Brazil was supported by the Ministry of Health of Brazil and the CNPq, grant 404067/2012-3. The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication. The authors declare that they have no competing interests.

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Plasmodium vivax infection compromises reticulocyte stability.

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